Cyanobacteria & Diatoms: Salinity Gradient Study

Environmental Microbiology (2000) 2(2), 217±226
Matching molecular diversity and ecophysiology of
benthic cyanobacteria and diatoms in communities
along a salinity gradient
Ulrich NuÈbel,1²* Ferran Garcia-Pichel,1³
Ester Clavero2 and Gerard Muyzer1§
1
Max Planck Institute for Marine Microbiology,
Bremen, Germany.
2
Departament QuõÂmica Ambiental, CID-CSIC,
Barcelona, Spain.
Summary
The phylogenetic diversity of oxygenic phototrophic
microorganisms in hypersaline microbial mats and
their distribution along a salinity gradient were investigated and compared with the halotolerances of closely
related cultivated strains. Segments of 16S rRNA genes
from cyanobacteria and diatom plastids were retrieved
from mat samples by DNA extraction and polymerase
chain reaction (PCR), and subsequently analysed by
denaturing gradient gel electrophoresis (DGGE). Sequence analyses of DNA from individual DGGE bands
suggested that the majority of these organisms was
related to cultivated strains at levels that had previously been demonstrated to correlate with characteristic salinity responses. Proportional abundances
of ampli®ed 16S rRNA gene segments from phylogenetic groupings of cyanobacteria and diatoms were
estimated by image analysis of DGGE gels and were
generally found to correspond to abundances of the
respective morphotypes determined by microscopic
analyses. The results indicated that diatoms accounted
for low proportions of cells throughout, that the cyanobacterium Microcoleus chthonoplastes and close relatives dominated the communities up to a salinity of
11% and that, at a salinity of 14%, the most abundant
cyanobacteria were related to highly halotolerant cultivated cyanobacteria, such as the recently established
phylogenetic clusters of Euhalothece and Halospirulina. Although these organisms in cultures had previously demonstrated their ability to grow with close
Received 5 October, 1999; revised 10 November, 1999; accepted 18
November, 1999. Present addresses: ²Montana State University,
Department of Land Resources and Environmental Sciences, 334
Leon Johnson Hall, Bozeman, MT 59717, USA. ³Arizona State University, Microbiology Department, Tempe, AZ, USA. §Netherlands
Institute for Sea Research (NIOZ), Den Burg (Texel), The Netherlands. *For correspondence. E-mail [email protected]; Tel.
(‡1) 406 994 3412; Fax (‡1) 406 994 4926.
Q 2000 Blackwell Science Ltd
to optimal rates over a wide range of salinities, their
occurrence in the ®eld was restricted to the highest
salinities investigated.
Introduction
It is a grand challenge for microbial ecologists to understand the distribution of microorganisms in nature in relation
to their physiological capabilities. Although their physiology
can be studied in great detail on strains growing in laboratory cultures, it has become increasingly apparent that
only a minority of the microorganisms occurring in nature
is represented by cultivated isolates. During the last decade, this insight has been promoted largely by the great
advances achieved in the detection of extant microbial diversity through the culture-independent analyses of nucleic
acids, particularly of small-subunit ribosomal RNAs and
the respective genes (for reviews, see Amann et al., 1995;
Pace, 1997; Muyzer, 1998; Ward et al., 1998). It is a common observation that 16S rRNA genes retrieved directly
from environmental samples rarely match those of cultivated strains or are only very distantly related to these
(e.g. Giovannoni et al., 1990; Ward et al., 1990; Barns et
al., 1994; Ferris et al., 1996; Hugenholtz et al., 1998).
Under such circumstances, it is not possible to infer the
organisms' ecologically relevant phenotypic properties
from their phylogenetic relationships. Consequently, the
distributions of gene clusters along environmental gradients are generally poorly understood in terms of the putative ecological niches of the underlying microorganisms
(Ruff-Roberts et al., 1994; Field et al., 1997; Massana et
al., 1997; Ward et al., 1998). In contrast, after previous isolation of representative cyanobacteria (Moore et al., 1998;
Urbach et al., 1998), the depth distribution of genetically
different oceanic picophytoplankton populations could
recently be explained in part by their differential adaptations to light intensity and spectral composition (Ferris
and Palenik, 1998; West and Scanlan, 1999).
We have attempted to unravel relationships between the
ecology and diversity of benthic cyanobacteria and diatoms
in hypersaline environments. In evaporation ponds of the
saltern in Guerrero Negro, Baja California, Mexico, the primary production of these microorganisms is the basis for
the existence of microbial mat ecosystems at salinities
from <6% to 16% (for reviews and detailed descriptions
218 U. NuÈbel, F. Garcia-Pichel, E. Clavero and G. Muyzer
of these sites, see Javor, 1989; Des Marais, 1995). Despite
the considerable interest among biogeochemists and microbiologists in these systems, until recently, only a limited
number of cyanobacterial and diatom isolates from hypersaline environments had been characterized physiologically and phylogenetically. During recent years, however,
this situation has changed to some extent. A signi®cant
number of clonal cultures has been recovered from microbial mats and characterized with respect to their phylogeny,
morphology, and physiology, the latter particularly concerning salinity responses (Table 1; Garcia-Pichel et al.,
1996; 1998; Karsten, 1996; E. Clavero et al., submitted;
NuÈbel et al., 2000). It remains to be demonstrated whether
the isolates now available adequately represent the organisms ¯ourishing in hypersaline mat ecosystems and if
the physiological data gained from laboratory cultures can
be con®dently extrapolated to ®eld conditions. As the comparative analysis of 16S rRNA gene sequences may offer
the means to examine the congruences or discrepancies
between strain collections and natural microbial communities, we recently developed a polymerase chain reaction
(PCR) protocol to amplify 16S rRNA gene segments
from cyanobacteria and from plastids of eukaryotic phototrophs from environmental samples (NuÈbel et al., 1997).
Here, we report on the application of this tool in combination with denaturing gradient gel electrophoresis (DGGE;
for a recent overview, see Muyzer, 1999) to investigate
the distribution of 16S rRNA genes from oxygenic phototrophic microorganisms in microbial mats along a gradient
of salinity. Data on the abundance of speci®c morphotypes
generally supported the results obtained.
Results
16S rRNA genes retrieved from microbial mats
Samples of microbial mats were collected along a gradient
of salinity in the saltern's evaporation ponds 2±6 (mats P2,
P3/4, P4, P5 and P6). Total nucleic acids were extracted
from mat samples before the ampli®cation of 16S rRNA
gene segments from cyanobacteria and diatom plastids
by PCR by applying phylum-speci®c primers (NuÈbel et
al., 1997). Sequence-dependent separation of the ampli®cation products by DGGE resulted in the band patterns
shown in Fig. 1. Fluorescence measurements and comparisons with a DNA mass calibration standard (Fig. 1) enabled
the calculation of amounts of DNA in individual gel bands.
DNA could be eluted, reampli®ed and sequenced (indicated in Fig. 1) from 29 out of 45 bands altogether. Bands
for which we were unable to obtain reliable sequences
each accounted for 0.5±4.4% (mean 1.9%) of the total
PCR products visible as ¯uorescent bands after DGGE.
Besides the generally low amounts of DNA forming
those bands, potential reasons for dif®culties with PCR
reampli®cations and subsequent sequence analyses
might have been insuf®cient separation of sequence-different DNA molecules by DGGE (e.g. bands P4.7, P6.6 and
P6.8). Similarly, the reampli®cation of DNA from bands
P2.7 and P2.8 resulted in PCR products migrating in subsequent DGGE at the position of band P2.10 (not shown),
a problem that is possibly caused by the reampli®cation of
incompletely focused predominant sequence populations
visible as `background smear' in the primary DGGE gel
(Fig. 1). However, we did not observe reampli®cation products forming patterns of bands in DGGE that would be
consistent with interpretation as the result of the generation of heteroduplex DNA molecules during PCR (Ferris
and Ward, 1997).
Nucleotide sequences obtained in this manner had a
length of 247±381 nucleotides, the latter corresponding to
the complete gene segment ampli®ed. Generally, the quality of the sequence data was high, with most sequences
containing no or single ambiguities (mean 2.4, maximum
14). These partial 16S rRNA gene sequences were suf®cient in all but ®ve cases to identify close relatives among
cultivated cyanobacteria and diatoms and could be robustly
integrated in the phylogenetic tree (Fig. 2). In all microbial
mats except that from evaporation pond 2, 16S rRNA gene
segments were found that were identical in sequence to
those from several Microcoleus chthonoplastes strains that
had recently been isolated from this and various other
habitats (Garcia-Pichel et al., 1996). In community P2,
Table 1. Halotolerances of cultivated cyanobacteria and diatoms from hypersaline environments.
Phylogeny of cultivated organisms
No. of strains
investigated
Halotolerance %
(optimum %)a
References
Lyngbya sp. MPI FGP-A
Microcoleus chthonoplastes
Euhalothece cluster
Halospirulina cluster
Oscillatoria limnetica
Diatoms
1
10
13
3
1
34
0±7 (0±3.5)
0±12 (0.5±9)
0±28 (1.5±25)
1.5±20 (3.5±20)
1.5±20 (6±15)
0.5±15 (0.5±12.5)
This study
Karsten (1996); Garcia-Pichel et al. (1996)
Garcia-Pichel et al. (1998)
NuÈbel et al. (2000)
Golubic (1980)
Clavero et al. (submitted)
a. Optimum indicates the salinity at which growth with at least half-maximal rate was measured for at least one member of the respective phylogenetic cluster.
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Matching molecular diversity and ecophysiology of benthic cyanobacteria
219
Fig. 1. Composite ®gure of ethidium bromide-stained DGGE separation patterns of PCR-ampli®ed segments of 16S rRNA genes. Results
derived from triplicate sampling cores are shown for each of the ®ve microbial mats (P2 to P6) investigated. Arrows indicate the positions and
designations of the DNA bands included in the subsequent analyses. Framed designations indicate bands from which nucleotide sequences
could be retrieved. Mixtures of PCR products derived from ®ve cyanobacterial strains were applied to each gel as standards [in lanes 1±3,
top to bottom, are Scytonema sp. strain B-77-Scy.jav., Synechococcus elongatus SAG 1402-1, Microcoleus chthonoplastes MPI-NDN-1,
Geitlerinema sp. strain PCC 9452 (`Microcoleus' sp. strain 10 mfx) and Euhalothece sp. strain PCC 7418]. The standard in lane 1 allows
gel-to-gel comparisons. The DNA mass calibration standard in lanes 2 and 3 enables the transformation of measured band ¯uorescence
values into amounts of DNA [in lane 2, the amounts of DNA in individual bands are (top to bottom) 528, 176, 59, 20 and 7 ng; in lane 3,
half of the amount of the mass calibration standard was applied.
two 16S rRNA gene sequences were found that each differed from the above sequences at single nucleotide positions. A number of sequences were detected that were
similar or identical, respectively (95.3±100%), to those
from cultivated cyanobacteria of the recently described
Euhalothece cluster (Garcia-Pichel et al., 1998). Other
sequences detected were highly similar (98.3±99.7%) to
those from cultivated cyanobacteria of the genera Halospirulina (NuÈbel et al., 2000) and Lyngbya (strain MPI
FGP-A was isolated from a microbial mat in the same
area) or from the strain Oscillatoria limnetica (originating
from hypersaline Solar Lake, Egypt; Cohen et al., 1975).
Several sequences were obviously derived from diatom
plastids. Only ®ve sequences could not be placed in
close af®liation to any oxygenic phototrophs for which
16S rRNA gene sequences have been determined to
date. Two of these as yet unidenti®able sequences (from
bands P4.9 and P5.7), however, clustered together signi®cantly, which could be con®rmed by calculating phylogenetic trees with various data subsets (not shown). In
mat P5, the PCR product derived from organisms of this
cluster amounted to 28% of the total ampli®ed DNA visible
in the DGGE bands. In all other mats, however, the relative
contribution of unidenti®ed sequences (including unsequenced bands) to the total was only minor, and 87% or
more of the total PCR products could be traced unequivocally to cyanobacteria and diatoms that are closely related
to known, cultivated strains. Although 16S rRNA gene
sequences not perfectly matching genes from isolates
were detected, our data suggest, in contrast to impressions left by previous studies (e.g. on hot spring mats),
that the cultures currently available represent well the
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phylogenetic diversity of oxygenic phototrophs present in
these hypersaline environments.
The distribution of 16S rRNA genes and
morphotypes among mats
Compared with other microorganisms, cyanobacteria and
diatoms display relatively complex cell and colony morphologies. This is the basis for their traditional classi®cation
(Anagnostidis and KomaÂrek, 1985; Castenholz and Waterbury, 1989; Round et al., 1990). The reliability of morphological criteria for the identi®cation of phylogenetically and
ecologically coherent taxa, however, may depend to a
large extent on the particular organisms of interest and
the habitats investigated. From the data available for
cyanobacteria, it appears that the simpler the morphology,
the more uncertain become predictions of phylogenetic
placement based on morphology (Ferris et al., 1996; Garcia-Pichel et al., 1996; NuÈbel et al., 2000). For well-studied
groups, however, microscopic observations may yield
valuable information about the distributions of these organisms in nature. For example, ®lamentous cyanobacteria
prevalent in many microbial mats the world over and
microscopically identi®able as Microcoleus chthonoplastes
were demonstrated to have virtually identical 16S rRNA
gene sequences (Garcia-Pichel et al., 1996). In contrast,
unicellular cyanobacteria and those with tightly, helically
coiled trichomes turned out to be phylogenetically very
diverse. However, in both these cases, basic morphology
together with the ability to thrive at high salt concentrations de®ned tight monophyletic clusters of organisms.
We termed the cluster Euhalothece (Garcia-Pichel et al.,
220 U. NuÈbel, F. Garcia-Pichel, E. Clavero and G. Muyzer
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Matching molecular diversity and ecophysiology of benthic cyanobacteria
221
Fig. 3. Proportional abundances of various
groups of oxygenic phototrophs in microbial
mats in relation to ambient salinity; note the
different scales on the ordinate axes of the
different graphs. Means and standard errors
from triplicate analyses are shown based on
cell counts (open circles) and amounts of
DNA measured in DGGE bands (closed
circles) respectively. Bars on top of each
graph illustrate the salinity tolerances for
the growth of cultivated strains of the
phylogenetic groups investigated (compare
Table 1). Black bars indicate growth with at
least half-maximal rates (optimal salinity
ranges); hatched bars indicate growth with
less than half-maximal rates (suboptimal
salinity ranges). Phylogeny- or morphologybased de®nitions of groups of organisms
investigated are given in the two columns
on the right. Numbers in brackets correspond
to morphotype illustrations in (NuÈbel et al.,
1999a). For taxon designations, see text. In
addition, the range of salinities is indicated
within which microbial mat development
occurs in the saltern system studied.
1998) and proposed the genus Halospirulina (formerly
classi®ed as Spirulina on the basis of morphology alone;
NuÈbel et al., 2000) respectively. Diatoms form a phylogenetically coherent group and are easily discernible
from cyanobacteria by light microscopy.
On the basis of these considerations, we can compare
light microscopic observations with analyses of 16S rRNA
genes from microbial mats. Figure 3 illustrates, in relation
to ambient salinities, the proportional abundances of morphological and phylogenetic groupings in the different mat
samples based on cell counts and amounts of DNA measured in DGGE bands respectively. Some technical limitations inherent to these quanti®cations, such as potential
PCR ampli®cation biases and overestimations of abundances of large cells, cannot be excluded here with certainty
and have been discussed previously (NuÈbel et al., 1999a).
It should be noted, however, that DGGE analysis is more
suitable to analyse the composition of PCR products quantitatively than the more widely applied molecular cloning
followed by subsequent random picking and screening of
transformants. Whereas in the latter approach, usually
less than a few hundred clones can be analysed, microscopic investigations of the (low diversity) oxygenic phototrophic mat communities studied here revealed that at least
2000±3000 individuals (cells) needed to be included in an
analysis to enable stable estimates of proportional abundances (NuÈbel et al., 1999a). DGGE band patterns in
each of the gel lanes in Fig. 1 are composed of <1012
DNA molecules and, thus, can be expected adequately
to re¯ect proportional abundances of sequence-different
Fig. 2. Maximum likelihood phylogenetic tree based on 16S rRNA gene sequences at least containing the nucleotides from 45 to 1455
(corresponding to the E. coli numbering; Brosius et al., 1981). 16S rRNA gene sequences from E. coli and Bacillus subtilis were used as
outgroup sequences. The phylogenetic af®liations of organisms represented by partial sequences (indicated by asterisks) were reconstructed
by applying the parsimony criteria without changing the overall tree topology (see ARB manual; Ludwig et al., 1998). Nucleotide sequences
derived from DGGE bands are indicated by band designations (compare Fig. 1), which are framed.
Q 2000 Blackwell Science Ltd, Environmental Microbiology, 2, 217±226
222 U. NuÈbel, F. Garcia-Pichel, E. Clavero and G. Muyzer
ampli®cation products (and presumably genes). A more
serious problem for the comparison of cell numbers with
gene abundances might be the varying number of 16S
rRNA genes per cell depending on the organisms' speci®c
identities. Microalgal plastids have been reported to contain
up to 650 copies of small circular genomes (Ersland et al.,
1981), whereas prokaryotes possess only 1±15 copies of
rRNA operons. Possibly, these differences in gene copy
numbers caused the proportional abundances of plastidderived PCR products to account for up to 36% of the
total, whereas in terms of cell numbers, diatoms were comparatively rare creatures, with proportional abundances of
maximally 4.7% and usually around or below 1% (Fig. 3).
Despite the potential drawbacks, abundance estimates
based on PCR and cell counts for the various groups of cyanobacteria were rather congruent, particularly for those
mat communities originating from higher ambient salinities. Both approaches indicated that Microcoleus chthonoplastes was the dominant inhabitant of all mats except
in pond 6, where the total salt concentration reached 14%.
Euhalothece-related 16S rRNA gene sequences were
found in mats from ponds 4, 5 and 6, with increasing abundance accompanying increasing salinity. These abundance
estimates coincided with cell counts of unicellular cyanobacteria. However, a second abundance maximum for
rod-shaped and spherical cyanobacteria was determined
microscopically at low salinity in mats P2 and P3/4. Most
probably, those organisms are marine forms and not
related to the highly halotolerant Euhalothece cluster.
Phylogenetic analyses suggest that these simplest cell
shapes evolved independently several times within the
cyanobacterial radiation (Turner, 1997), and the correlation of unicellular morphology and phylogenetic af®liation to Euhalothece will hold only in high-salinity habitats
(Garcia-Pichel et al., 1998). Halospirulina-related sequences were found in P6 exclusively, coinciding with the
detection of sheathless, helically coiled ®laments, which
is the characteristic appearance of these cyanobacteria
(NuÈbel et al., 2000). Similarly, the only sequence derived
from an organism related to the isolate Oscillatoria limnetica (Phormidium hypolimneticum sensu Campbell,
1985) was detected at 14% salinity; ®laments morphologically resembling this strain were abundant in this mat, but
rare or undetectable in all other samples. Sequences
related to Lyngbya strains (sensu Castenholz, 1989)
were found in mat P2 exclusively. This distribution is
consistent with the microscopic detection of Lyngbya-like
®laments.
Discussion
The results presented suggest that the phylogenetic diversity of oxygenic phototrophic microorganisms dominating
the variety of microbial mats investigated along the salinity
gradient is well represented by cultivated cyanobacteria
and diatoms whose ecophysiology is known. In the samples
originating from hypersaline brines (above 7% total salts;
Por, 1980), we were able to identify the majority of sequences as being derived from organisms closely related
to groups of isolated strains. Within established phylogenetic lineages, those 16S rRNA gene sequences that do
not perfectly, but only closely, match previously known
gene sequences indicate the extent to which the diversity
in the mats may exceed the diversity currently cultivated.
The sequence divergence detected amounted to maximally 4.7% in the Euhalothece cluster, 5.6% in the clade
of diatom plastids and less than 0.6% in the remaining
clusters. However, for none of the clusters did the gene
segments detected in the mats increase the maximum
sequence divergence as established on the basis of cultivated strains. This was found despite the fact that the divergence of corresponding full-length 16S rRNA genes gets
slightly overestimated if the comparison is limited to the
partial segments analysed here. Because rRNA genes are
highly conserved in function and structure, the sequence
divergence observed may still correspond to signi®cant,
yet unknown, phenotypic differences among the underlying organisms (Palys et al., 1997). However, for the clusters involved in this study, a remarkable ecophysiological
uniformity in their salinity tolerance has been described
(Karsten, 1996; Garcia-Pichel et al., 1996; 1998; E. Clavero
et al., submitted; NuÈbel et al., 2000), indicating that the
adaptation to life in high-salinity waters is an ancient and
conserved evolutionary development that probably has
been acquired by several lineages independently (GarciaPichel et al., 1998; NuÈbel et al., 2000). Therefore, the
cultures studied are most probably representative of the
physiological tolerances to salinity of the majority of mat
community components (Table 1). Isolates of Microcoleus
chthonoplastes and diatoms with various generic and speci®c identities were somewhat variable regarding their
halotolerances, corresponding to the habitats that they
had been isolated from. Members of both groups, however,
commonly preferred marine salinity and grew only poorly or
not at all at total salt concentrations higher than 12% (Fig.
3; Karsten, 1996; E. Clavero et al., submitted). Cultivated
strains of extremely halotolerant unicellular cyanobacteria
formed one or possibly two phylogenetic clusters (including
the monophyletic Euhalothece cluster), as indicated
by their 16S rRNA gene sequences. Similarly, highly halotolerant cyanobacteria with helically coiled trichomes
(Halospirulina spp.) are closely related to each other and
distinct from their morphological counterparts in freshwater
and temperate marine habitats. In addition, isolates representing these two latter groups can be termed extremely
euryhaline, because they are able to grow with close to
optimal rates over wide ranges of salinities (Table 1; Garcia-Pichel et al., 1998; NuÈbel et al., 2000). The phylogeny
Q 2000 Blackwell Science Ltd, Environmental Microbiology, 2, 217±226
Matching molecular diversity and ecophysiology of benthic cyanobacteria
and halotolerances of relatives of Lyngbya sp. and Oscillatoria limnetica still await systematic investigations. However, we tentatively include here some preliminary data
on these cyanobacteria (Table 1), because we detected
related gene sequences and the respective morphotypes
in some of the mat samples.
In terms of salinity, the fundamental ecological niches of
cyanobacteria and diatoms of the different groups overlap
greatly (Table 1). However, the actual distribution of these
organisms in the natural environment is much more
restricted than dictated by their speci®c physiological tolerances to salinity (Fig. 3). The development of cyanobacterial mats at salinities below 6% is limited by competition with
larger eukaryotic algae and higher plants (Des Marais,
1995) and increased grazing pressure (Javor and Castenholz, 1984). Gypsum precipitation, starting at a salinity of
<16%, may prevent nutrient recycling from sediments and
thereby demarcate an upper salinity limit for microbial mat
maintenance (Javor, 1989). Within this range, the mat communities investigated differed in composition along the
salinity gradient (Fig. 3). Most remarkably, perhaps, the
highly euryhaline cyanobacteria (Euhalothece, Halospirulina and, tentatively included here, O. limnetica ) are abundant in the most saline brines only. This restriction is
probably the result of interactions with hostile organisms,
such as competitors, predators or parasites, the occurrence
of which may again correlate with salinity. M. chthonoplastes dominated the mats up to 11% total salt concentration, above which its proportional abundance greatly
decreased, coinciding with the upper limit for growth of
related cultivated strains (Karsten, 1996). The reasons
for the superiority of M. chthonoplastes are still little understood, however. Ecological niches must be considered
multidimensional, and organisms that occupy overlapping
(fundamental or realized) niches along one dimension,
such as salinity, can be expected to differ with respect to
some other property, such as their adaptations to light or
nutrient availability or resistances to hostile substances
or neighbours. Complex interactions of cyanobacteria,
invertebrate grazers and environmental gradients have
been described for microbial mats in hot springs (Wickstrom and Castenholz, 1985) and may also be expected
here. In any case, our results indicate that the (post-interactive) distributions of cyanobacteria in nature in comparison with their physiological tolerances may be rather
restricted. These realized niches (Begon et al., 1990) do
not necessarily centre around the respective salinity
optima as has been hypothesized previously (Golubic,
1980). Highly halotolerant cyanobacteria acclimate to elevated ambient salinity by synthesizing quaternary ammonium compounds that function as osmolites, but they keep
the physiological potential to thrive at lower salinities
(MacKay et al., 1984; Reed and Stewart, 1988). Possibly,
such a pattern of adaptations and distributions is rather
Q 2000 Blackwell Science Ltd, Environmental Microbiology, 2, 217±226
223
common among microorganisms that have gained the
ability to tolerate hostile environmental conditions through
evolution.
The remarkable match of cultivation-based and molecular
biological samplings of microbial diversity reported here is
in part certainly caused by restriction of the study to a
single functional group of organisms. In addition, the
morphologies of cyanobacteria and diatoms in the habitats
investigated to some extent correlate with their phylogeny
and, thus, microscopic examinations aid in the identi®cation of organisms yet to be cultivated. This contrasts with
the situation in microbial mats found in thermal environments, from which, until recently, only a single type of
unicellular cyanobacterium had been isolated in culture,
simply because the cyanobacterial diversity present had
long been overlooked as a result of uniform morphology
(Ferris et al., 1996). This ®nding, however, suggests that
the commonly found discrepancies between the compositions of strain collections and natural microbial communities
may arise to a considerable extent from researchers'
inability or neglect in recognizing unique and predominant
microorganisms (Pinhassi et al., 1997), and not necessarily from their apparent immodesty or `unculturability'.
However, to appreciate fully the diversity of microorganisms, characteristics other than the highly conserved
16S rRNA genes need to be analysed (Palys et al.,
1997; Ward, 1998). Future research is required towards
a more comprehensive understanding of the complex
interactions of multiple physical, chemical and biological
factors that determine the composition of these microbial
mat communities.
Experimental procedures
Sampling
Sampling sites were located in evaporation ponds of the saltern in Guerrero Negro, Baja California Sur, Mexico (Javor,
1989; Des Marais, 1995). Microbial mats were sampled in
April 1996 (mats P2, P4 and P6) and April 1997 (P3/4 and
P5). Mat designations correspond to designations of evaporation ponds 2±6, along which a gradient of salinity is maintained that ranges from 5.5% to 14% total salt concentration
(Fig. 3; NuÈbel et al., 1999b). Macroscopic characteristics of
the mats investigated and ®eld conditions have been described
previously (NuÈbel et al., 1999a). For the isolation of Lyngbya
sp., pieces from an intertidal mat were sampled in the salt
marsh adjacent to the saltern, air dried and transported to
the laboratory at room temperature. For microscopic and molecular biological investigations, triplicate cores 10±20 cm apart
were taken as samples from each mat and analysed independently. For light microscopy, mat samples were ®xed in 5%
formaldehyde and stored at 48C. After longer storage of samples (several months), this preservation method resulted in
slight shrinking of cyanobacterial cells that did not, however,
prevent their proper identi®cation. For nucleic acid extractions,
224 U. NuÈbel, F. Garcia-Pichel, E. Clavero and G. Muyzer
the mat samples were frozen on site, transported to the laboratory in liquid nitrogen and stored at 80 8C until processed.
Subsequent analyses were restricted to the photic zones of
each of the mats, de®ned by the maximum depths at which
gross photosynthesis could be detected with oxygen microelectrodes in a ®eld laboratory (Garcia-Pichel et al., 1999).
Microscopy
The layers corresponding to the photic zones were cut from
formaldehyde-®xed mat samples with scalpel blades. Pieces
representing <0.5 ´ 0.5 mm mat surface area were placed
on glass slides and chopped and stirred to achieve even distribution. Randomly chosen phase-contrast microscopic ®elds
were investigated at 400-fold magni®cation, and 2000±3000
cells of cyanobacteria and diatoms per sample were counted
to ensure representative results (NuÈbel et al., 1999a). Green
non-sulphur bacteria (Chloro¯exaceae ) were distinguished
from cyanobacteria by their lack of visible ¯uorescence and
omitted from the analyses.
Molecular biological techniques
DNA extraction, PCR and DGGE have been described previously (NuÈbel et al., 1997; 1999a). Brie¯y, the layers corresponding to photic zones were cut aseptically from mat cores
(representing <60 mm2 of mat surface) and homogenized in
Dounce tissue homogenizers (Novodirect). The suspensions
were repeatedly frozen and thawed and subsequently incubated in the presence of SDS and proteinase K. Cell lysis
was controlled microscopically. DNA was extracted by applying hexadecyltrimethylammonium bromide, phenol, chloroform and isoamyl alcohol and precipitated by the addition of
isopropyl alcohol. The oligonucleotide primers CYA359F and
CYA781R were applied to amplify 16S rRNA gene segments
selectively from cyanobacteria and plastids (NuÈbel et al.,
1997). The numbers in the primer designations refer to the
58 ends of target signature sites in 16S rRNA genes (Escherichia coli nucleotide numbering; Brosius et al., 1981). A 40
nucleotide GC-rich sequence was attached to the 58 end of
the primer CYA359F to improve the detection of sequence
variation in ampli®ed DNA fragments by subsequent DGGE.
As templates for ampli®cations, 10 ng of DNAs extracted
from mat samples was added to each 100 ml reaction mixture.
Ampli®ed DNA (500 ng) was applied to denaturing gradient
gels. For DGGE, polyacrylamide gels with a denaturant gradient from 20% to 60% were used, and electrophoreses were
run for 3.5 h at 200 V and 60 8C. Ethidium bromide-stained
gels were irradiated with UV light and photographed with a
digital image gel documentation system (Cybertech). The
intensities of gel band ¯uorescences were measured on digital
images by applying the gel-plotting macro implemented in the
NIH IMAGE software package, version 1.62 (National Institutes
of Health). Fluorescence values of individual bands were transformed into amounts of DNA by comparison with a DNA mass
calibration standard for DGGE (Fig. 1; NuÈbel et al., 1999a). To
determine nucleotide sequences of DNA molecules forming
individual bands in DGGE, the bands were cut from the gels
and incubated in 50 ml of TE buffer (10 mM Tris-HCl pH 8.0,
1 mM EDTA) at 48C for elution of DNA. Incubation times varied
from 3 to 20 days with no noticeable effect on the results. These
solutions (1 ml) were applied as templates in PCRs applying
the primers CYA359F and CYA781R. The resulting reampli®cation products were checked by DGGE analysis for correspondence to the bands of interest, puri®ed by applying the
QIAquick PCR puri®cation kit (Qiagen) and sequenced by
applying the Applied Biosystems PRISM dye terminator
cycle sequencing ready reaction kit and 377 DNA sequencer.
For the majority of bands, the sequences of both DNA strands
were determined using the same primers as those used for
ampli®cation, while omitting the GC clamp from primer
CYA359F.
The EMBL accession numbers for the 16S rRNA gene sequences determined in this study are AJ248289±AJ248317.
Phylogeny reconstruction
Cyanobacterial and plastid 16S rRNA gene sequences available from GenBank were aligned to the sequences in the database of the software package ARB, developed by W. Ludwig and
O. Strunk, and available at http:/ /www.mikro.biologie.tumuenchen.de. A phylogenetic tree was constructed on the
basis of almost complete sequences (from nucleotide positions
45 to 1455 corresponding to the E. coli numbering) by applying the maximum likelihood method as integrated in the ARB
software package. Alignment positions at which one or more
sequences had gaps or ambiguities were omitted from the
analyses. Partial sequences (including those representing
DGGE bands) were integrated in the dendrogram according
to the maximum parsimony criterion without allowing them
to change the topology of the tree as established with complete
sequences (see ARB manual; Ludwig et al., 1998).
Isolation and cultivation of Lyngbya sp.
Sea water and hypersaline media were prepared by dissolving
appropriate amounts of commercial sea water salts mixture in
distilled water to which nutrients, trace elements and vitamins
were added according to Provasoli's enriched sea water formulation (Starr and Zeikus, 1987). Freshwater medium was
BG11 (Rippka, 1988). Solid media contained 1% agarose.
Cultures received constant irradiance of 20 mmol photons
m 2 s 1 of ¯uorescent light. Cyanobacteria morphologically
resembling Lyngbya sp. (Castenholz, 1989) were isolated
from an intertidal microbial mat that is exposed to desiccation
periodically. Clonal strains were gained by repeatedly transferring individual ®laments after their self-isolation through
gliding movements on the surface of solid medium (Rippka,
1988). The halotolerance of strain MPI FGP-A was determined
by visual inspection of growth in test tube cultures with liquid
media. Salinities tested were 0%, 1.5%, 3.5%, 7% and 10%.
Acknowledgements
We gratefully acknowledge Exportadora de Sal, S.A. de C.V.,
Mexico, for allowing us to conduct research on their premises
and for continued logistic support, and we thank two anonymous
reviewers for helpful comments. This work was supported ®nancially by the Max Planck Society and the Deutsche Forschungsgemeinschaft, Germany. E.C. received a fellowship from the
Ministerio de EducacioÂn y Ciencia, Spain (FPI94).
Q 2000 Blackwell Science Ltd, Environmental Microbiology, 2, 217±226
Matching molecular diversity and ecophysiology of benthic cyanobacteria
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